Homers are scaffolding proteins that bind G protein–coupled receptors (GPCRs), inositol 1,4,5-triphosphate (IP3) receptors (IP3Rs), ryanodine receptors, and TRP channels. However, their role in Ca2+ signaling in vivo is not known. Characterization of Ca2+ signaling in pancreatic acinar cells from Homer2−/− and Homer3−/− mice showed that Homer 3 has no discernible role in Ca2+ signaling in these cells. In contrast, we found that Homer 2 tunes intensity of Ca2+ signaling by GPCRs to regulate the frequency of [Ca2+]i oscillations. Thus, deletion of Homer 2 increased stimulus intensity by increasing the potency for agonists acting on various GPCRs to activate PLCβ and evoke Ca2+ release and oscillations. This was not due to aberrant localization of IP3Rs in cellular microdomains or IP3R channel activity. Rather, deletion of Homer 2 reduced the effectiveness of exogenous regulators of G proteins signaling proteins (RGS) to inhibit Ca2+ signaling in vivo. Moreover, Homer 2 preferentially bound to PLCβ in pancreatic acini and brain extracts and stimulated GAP activity of RGS4 and of PLCβ in an in vitro reconstitution system, with minimal effect on PLCβ-mediated PIP2 hydrolysis. These findings describe a novel, unexpected function of Homer proteins, demonstrate that RGS proteins and PLCβ GAP activities are regulated functions, and provide a molecular mechanism for tuning signal intensity generated by GPCRs and, thus, the characteristics of [Ca2+]i oscillations.
Receptor-mediated Ca2+ signaling consists of a biochemical component that hydrolyzes phosphatidylinositol-bisphosphate (PIP2)*
Abbreviations used in this paper: BS, bombesin; CPA, cyclopiazonic acid; GAP, GTPase-activating protein; GPCR, G protein–coupled receptors; IP3, inositol 1,4,5-triphosphate; IP3R, IP3 receptor; pAb, polyclonal antibody; PIP2, phosphatidylinositol-bisphosphate; PMCA, plasma membrane Ca2+ ATPase; RGS, regulators of G proteins signaling; SERCA, sarco/endoplasmic reticulum Ca2+ ATPase; SLO, streptolysin O; WT, wild-type.
The highly coordinated [Ca2+]i oscillations and waves require polarized expression of Ca2+-signaling proteins, their organization into complexes, and regulation of each component within the signaling complex. Indeed, Ca2+-signaling proteins are clustered in microdomains of polarized cells, such as the pre- and postsynaptic membranes in neurons (Hering and Sheng, 2001) and the apical pole of secretory cells (Kiselyov et al., 2003). Signaling complexes are assembled with the aid of scaffolding proteins that express multiple protein–protein interacting domains (Hering and Sheng, 2001; Minke and Cook, 2002). The role of scaffolding proteins in tyrosine kinase receptors (Hunter, 2000) and cAMP/PKA-mediated signaling (Smith and Scott, 2002) is well characterized. Much less is known about scaffolding proteins in Ca2+ signaling. In synapses, PSD-95, SHANK, GRIP, and probably other scaffolds, participate in assembly of signaling complexes, including Ca2+ signaling (Hering and Sheng, 2001). InaD is the scaffold that assembles Ca2+-signaling complexes in Drosophila photoreceptors (Minke and Cook, 2002). However, the primary scaffolding protein that assembles Ca2+-signaling complexes in nonneuronal cells is not known. Homer proteins have recently emerged as attractive candidates (Fagni et al., 2002). Homers are scaffolding proteins that are composed of an EVH protein–binding domain, a coiled-coil multimerization domain, and a leucine zipper (Fagni et al., 2002). The EVH domain binds the GPCR mGluR1α/5, IP3Rs, ryanodine receptors, and probably other proteins involved in Ca2+ signaling (Tu et al., 1998; Xiao et al., 1998, 2000). However, the present work reveals that Homers may not function as simple scaffolds, as deletion of Homer 2 or 3 did not disrupt polarized localization of IP3Rs and other Ca2+-signaling proteins in pancreatic acini, but rather affected the efficiency of signal transduction.
G proteins amplify and transduce signals from the receptor to the appropriate effector, and are, thus, a central regulatory site of signal transduction efficiency. Activation of G proteins involves a receptor-catalyzed GDP-GTP exchange reaction on the α subunit to release Gα·GTP and Gβγ (Gilman, 1987), which, in turn, activate separate effector proteins (Gudermann et al., 1997). The off reaction entails the hydrolysis of GTP and reassembly of the Gα·GDPβγ heterotrimer. This reaction is accelerated by two separate GTPase-activating proteins (GAPs), the PLCβ effector protein (Ross, 1995) and the regulators of G proteins signaling (RGS) proteins (Ross and Wilkie, 2000). In vitro (Ross and Wilkie, 2000) and in vivo studies (Cook et al., 2000) suggest that both catalytic mechanisms participate in Ca2+ signaling. Furthermore, regulation of Gαq by RGS proteins confers receptor-specific Ca2+ signaling (Xu et al., 1999), drives [Ca2+]i oscillations (Luo et al., 2001), and probably accounts for the oscillation in [IP3] during [Ca2+]i oscillations (Hirose et al., 1999; Nash et al., 2001). [Ca2+]i oscillations due to [IP3] oscillations require cyclical activation and inactivation of RGS and/or PLCβ GAP activity. To date, little is known about the regulation of RGS proteins and PLCβ GAP activity.
The results reported here show that Homer 3 does not have a major role in Ca2+ signaling in pancreatic acinar cells, whereas Homer 2 regulates GAP activity of both RGS proteins and PLCβ. Deletion of Homer 2 resulted in increased potency of agonist-stimulated Ca2+ signaling and, thus, increased frequency of agonist-evoked [Ca2+]i oscillations. This phenotype was traced to reduced GAP activity of exogenous RGS proteins to inhibit Ca2+ signaling in cells from Homer2−/− mice. In support of this mechanism, Homer 2 preferentially binds to PLCβ and activates RGS4 and PLCβ GAP activity in an in vitro reconstitution system. Thus, Homer 2 tunes the intensity of Ca2+ signaling to regulate [Ca2+]i oscillation frequency and, thus, cell functions regulated by this signaling pathway.
Homer proteins are encoded by three genes, Homer 1, 2, and 3 (Xiao et al., 2000; Fagni et al., 2002). Involvement of Homers in Ca2+ signaling was concluded from binding of GPCRs and IP3Rs to the EVH domain of Homer and the partial inhibition of Ca2+ signaling in neurons by expression of Homer 1a, a natural dominant negative of Homer scaffolding function (Tu et al., 1998). Based on these observations, it is generally assumed that Homers function as scaffolds that assemble and retain Ca2+-signaling complexes in cellular microdomains (Hering and Sheng, 2001; Fagni et al., 2002). To probe the role of Homers in Ca2+ signaling more directly, we generated mice deficient in specific Homer isoforms and characterized Ca2+ signaling in cells from these mice. We focused on pancreatic acinar cells because these polarized cells express all Ca2+-signaling proteins in a highly polarized manner at the apical pole (Lee et al., 1997a,b) and generate Ca2+ signals in the form of propagating Ca2+ waves (Kasai et al., 1993; Shin et al., 2001).
Deletion of Homers does not affect polarized expression of IP3Rs
The most basic function of a scaffolding protein is targeting and retention of its binding partners in a defined microdomain. Therefore, we first compared localization and expression of the Homers and IP3Rs in cells from wild-type (WT) and mutant mice. Fig. 1
describes the generation of the Homer 2−/− mice and shows that the gene was deleted (B), and the mRNA (B) and protein (C) could not be detected. Fig. 2
illustrates the localization of the three Homer isoforms and of IP3Rs in cells from WT and Homer2−/− mice. It is evident that Homers 1 and 2 are expressed exclusively in the apical pole of pancreatic acinar cells (Fig. 2, A–D), the site enriched in all three isoforms of IP3Rs (Fig. 2, G–L) and other Ca2+-signaling proteins (Lee et al., 1997a,b; Shin et al., 2001; Zhao et al., 2001) and from which Ca2+ waves emanate (Xu et al., 1996a; Shin et al., 2001). On the other hand, Homer 3 is expressed in the basal pole (Fig. 2, E and F). Deletion of Homer 2 did not affect expression of Homer 1 or 3. Most notably, deletion of Homer 2 had no effect on the localization of any IP3R isoform. This was probably not due to the compensatory effect of other Homers because preliminary experiments with pancreatic acini from one mouse from which Homers 1, 2, and 3 were deleted showed no obvious effect of Homers deletion on IP3Rs localization. This was unexpected in view of the binding of IP3Rs to the EVH domain of Homers (Tu et al., 1998).
Next, we examined whether deletion of Homer 2 affected expression of the Ca2+ transporters that generate the Ca2+ signal (Fig. 3)
. Although all proteins could be detected, attempts to quantify protein expression by Western blot using membranes prepared from pancreatic acini were largely unsuccessful due to excessive protein degradation. As reported before, this was particularly the case for all IP3Rs (Lee et al., 1997b). Reliable results could be obtained only for SERCA2b, the expression level of which increased 2.2-fold in pancreatic acinar cells from Homer 2-deficient mice (Fig. 3 A). Similar results were obtained with parotid gland (not depicted) and brain membranes (Fig. 3 B). The immunolocalization in Fig. 2 suggests that deletion of Homer 2 has minimal or no effect on expression of all IP3R isoforms. This was further tested in brain extracts. For the most part, the levels of IP3Rs were unaffected, although a slight increase in IP3R1 may have occurred in Homer2−/− cells (Fig. 3, C and D). Similarly, expression of PMCA was unaltered in Homer2−/− cells (Fig. 3 E). These findings imply that the expression and retention of IP3Rs in cellular microdomains do not require the expression of Homer 2. This conclusion can probably be extended to other components of the Ca2+-signaling complex.
Homer 3 does not participate in Ca2+ signaling in secretory acinar cells
Expression of Homer 3 at the basal pole of pancreatic acinar cells, its absence from the apical pole, and the specificity of Homer 3 localization is further illustrated in Fig. 4
(A and B). A comparison of Ca2+ signals obtained in WT and Homer 3−/− cells stimulated with high agonist concentration to trigger a single [Ca2+]i transient (Fig. 4 C) or stimulated with low agonist concentration to evoke [Ca2+]i oscillations (Fig. 4 D) revealed no clear difference in any parameter of the Ca2+ signals. Likewise, no differences were observed in the Ca2+ signals recorded from WT and Homer 3−/− cells using the protocols in Figs. 4 and 5
. Deletion of Homer 3 also had no effect on localization of Ca2+-signaling proteins in the apical pole of pancreatic, submandibular, and parotid gland acinar and duct cells, as was found for Homer 2−/− (unpublished data). Hence, it appears that Homer 3 does not participate directly in controlling GPCR-dependent Ca2+ signaling in the cells and tissue types examined. This is not surprising considering the differential localization of Ca2+-signaling complexes and Homer 3. The role of Homer 3 in cell function remains to be defined. Preliminary experiments with cells from Homer1−/− mice showed that deletion of Homer 1 did not affect agonist-stimulated IP3 production or Ca2+ mobilization in pancreatic acini (unpublished data), a phenotype different from that of cells from Homer 2−/− mice.
Deletion of Homer 2 increases potency of agonist stimulation
A different picture emerged when Ca2+ signaling was analyzed in cells from Homer2−/− mice. The effect of Homer 2 deletion on Ca2+ signaling was first studied in maximally stimulated WT and Homer2−/− acinar cells. Fig. 5 A and the summary in Fig. 5 C show that 1 mM carbachol increased [Ca2+]i to a higher level in Homer2−/− cells. In addition, Homer2−/− cells stimulated with carbachol reduced [Ca2+]i to a stable plateau faster than cells from WT mice. However, the plateau level was similar in both cell types. The higher [Ca2+]i increase in Homer2−/− cells could be due to increased Ca2+ release from internal stores and/or increased Ca2+ influx across the PMCA. To determine the contribution of each pathway we compared Ca2+ influx in the two cell types by a Ca2+-removal and -readdition protocol in stimulated cells. The cells were also treated with the SERCA pump inhibitor cyclopiazonic acid (CPA) to maximally deplete the stores and activate Ca2+ entry. Fig. 5 A (inset) shows that deletion of Homer 2 had no effect on Ca2+ influx. Similar results were observed in all seven similar experiments. Furthermore, removal of Ca2+ from the medium and stimulation with carbachol and CPA did not reduce the difference in [Ca2+]i increase between the two cell types (Fig. 5 C, middle columns, results summary). This indicates a higher Ca2+ load of the ER of Homer2−/− cells, as expected from the increased expression of SERCA2b demonstrated in Fig. 3. Therefore, the higher increase in [Ca2+]i in Homer2−/− cells is due to increased Ca2+ release from internal stores. The faster rate of [Ca2+]i decrease after Ca2+ release can be due to faster pumping of the Ca2+ out of the cells by PMCA or due to faster clearance of cytosolic Ca2+ by SERCA pumps. This was addressed by inhibiting SERCA pump activity at the onset of cell stimulation. Fig. 5 B shows that under these conditions [Ca2+]i was still increased to a higher level in Homer2−/− cells, but now the rate of [Ca2+]i reduction was similar in WT and Homer2−/− cells. In summary, the observed altered properties of the Ca2+ signal in maximally stimulated Homer2−/− cells is precisely what is expected from the adaptive increased expression of SERCA2b shown in Fig. 3.
To probe Ca2+ signaling further, we examined the response to increasing concentrations of carbachol. At low concentrations, agonists evoke [Ca2+]i oscillations. The frequency, and in some cases, the amplitude, of the oscillations increases with increased agonist concentration until at a high enough agonist concentration the oscillations merge into a single transient increase in [Ca2+]i (Berridge, 1993). This pattern is shown in Fig. 5 (D–F), left traces, for cells from WT mice. At 1 μM, carbachol induced low frequency oscillations (see Fig. 6 B for summary). The residual Ca2+ content in the stores was estimated by discharging it by exposing the cells to 1 mM carbachol and 10 μM CPA. 2.5 μM carbachol caused a substantial initial increase in [Ca2+]i that was followed by high frequency–low amplitude oscillations. Finally, 10 μM carbachol evoked a large increase in [Ca2+]i with high frequency oscillations superimposed on the downward stroke of [Ca2+]i. 10 μM carbachol released ∼70% of stored Ca2+. Remarkably, deletion of Homer 2 increased the response at all carbachol concentrations between 1 and 10 μM. Thus, 1 μM carbachol induced a response in Homer2−/− cells similar to that induced by 2.5 μM carbachol in WT cells. 2.5 μM carbachol caused a transient increase in [Ca2+]i while releasing ∼60% of stored Ca2+, whereas 10 μM carbachol mobilized the entire intracellular Ca2+ pool of Homer2−/− cells, similar to the effect of 100 μM carbachol in WT cells.
Deletion of Homer 2 increased the potency of all Gq-coupled receptors expressed in acinar cells examined. Fig. 6
shows part of the results obtained with bombesin (BS) and CCK stimulation, focusing on the physiological response of [Ca2+]i oscillations. CCK at 20 pM induced typical baseline [Ca2+]i oscillations in cells from WT mice. The [Ca2+]i oscillations evoked by the same concentration of CCK in cells from Homer2−/− mice occurred at a frequency about twice as higher than those recorded in cells from WT mice (Fig. 6, A and B). The traces in Fig. 6 C show that increasing BS concentration from 50 to 100 pM increased the frequency of the oscillations by 1.7-fold in WT cells. The same fold increase in frequency (1.8) was observed in Homer2−/− cells, but at each BS concentration, the frequency of the oscillations was ∼2.3-fold higher in Homer2−/− cells. In aggregate, the results of Figs. 5 and 6 show that deletion of Homer 2 increases the potency of agonists to stimulate Ca2+ signaling by GPCRs. The increase in potency is due to a change in a regulatory step common to signaling by GPCRs because the response evoked by all GPCRs examined was affected. In the next stage, we searched for this Homer 2–regulated general mechanism.
Homer 2 does not affect IP3-mediated Ca2+ release
The frequency of Ca2+ oscillations is controlled by a biochemical mechanism through the production of IP3 (Luo et al., 2001) and by several biophysical mechanisms that control the activity of the IP3Rs (Kiselyov et al., 2003). The activity of IP3Rs is regulated by the concentrations of IP3 and [Ca2+]i (Thrower et al., 2001), as well as Ca2+ content in the stores, which increases channel sensitivity to IP3 (Missiaen et al., 1992; Xu et al., 1996b). Because of the increased SERCA2b and Ca2+ content in the stores of Homer2−/− cells, we considered the possibility that Ca2+ release from the stores of these cells was more sensitive to IP3, leading to increased frequency of [Ca2+]i oscillations. The results in Fig. 7
exclude this possibility. Fig. 7 (A–C) shows the same potency for IP3 to release Ca2+ from the stores of streptolysin O (SLO)–permeabilized WT and Homer2−/− cells. In Fig. 7 (D–H), the frequency of IP3-evoked [Ca2+]i oscillations was measured in intact cells. In these experiments, the whole cell mode of the patch clamp technique was used to record the Ca2+-activated Cl− current as a reporter of [Ca2+]i (Zeng et al., 1996). The controls in Fig. 7 (D and F) show that [Ca2+]i oscillations evoked by 0.5 μM carbachol occurred at a higher frequency (1.86- ± 0.26-fold, n = 7) in Homer2−/− compared with control cells, whereas the response to maximal agonist stimulation was the same in both cell types. Long-lasting, IP3-evoked [Ca2+]i oscillations were induced by infusing the cells with 20 μM of the nonhydrolyzable IP3 analogue, 2,4,5 IP3. Fig. 7 (E–G) shows that IP3 evoked [Ca2+]i oscillations with indistinguishable frequency in the two cell types. The implications of the findings in Fig. 7 are that the function of the IP3Rs is the same in cells from WT and Homer2−/− mice and that the increased GPCR sensitivity to agonist stimulation in Homer2−/− cells is due to regulation by Homer 2 of a step upstream of Ca2+ release by IP3.
Homer 2 regulates RGS proteins and PLCβ GAP activities
Enhanced activity of the biochemical component of Ca2+ signaling in Homer2−/− cells implies enhanced activation of PLCβ and IP3 production, which can account for the increased signal intensity and frequency of Ca2+ oscillations. This was tested by measuring the dose response for carbachol-stimulated IP3 production. Notably, Fig. 8
A shows that deletion of Homer 2 increased the potency of carbachol in stimulating IP3 production about fivefold without affecting maximally stimulated IP3 production. The same phenomenon was observed using two submaximal and one maximal concentration of CCK and BS (unpublished data).
IP3 production entails activation of Gq by receptors and activation of PLCβ by Gαq. The steady-state level of active Gαq is determined by the balance between the receptor-catalyzed GTP-GDP exchange reaction on Gαq and hydrolysis of GTP by Gαq (Gilman, 1987). GTP hydrolysis is accelerated by RGS proteins (Ross and Wilkie, 2000) and by the effector itself, PLCβ (Ross, 1995). The increased potency of agonists in Homer2−/− cells could be due to increased activation of PLCβ by Gαq-GTP, stimulation of the receptor catalyzed GTP-GDP exchange reaction, inhibition of the GAP activity of RGS proteins and PLCβ, or any combination of the above. The first possibility was tested by measuring the dose response for activation of PLCβ by GTPγS. Fig. 8 B shows that activation of PLCβ by GTPγS (activated Gαq) was the same in WT and Homer2−/− cells. Hence, deletion of Homer 2 did not affect activation of PLCβ by Gαq.
To test the second possibility, we measured the inhibition of Ca2+ signaling by RGS4, which was infused into the cells using a patch pipette. Due to expression of several RGS proteins in one cell and the lack of information as to the specific RGS protein in each Ca2+-signaling complex, it was not possible to manipulate the native RGS proteins activity. Instead, we measured the ability of exogenous RGS4 to inhibit Ca2+ signaling. Fig. 9
(A–C) shows that in WT cells, 0.25 nM RGS4 abolished [Ca2+]i oscillations in response to 0.5 μM carbachol and inhibited >90% of the response to maximal stimulation with 1 mM carbachol. In contrast, in Homer2−/− cells, 0.25 nM RGS4 only partially inhibited the response to maximal carbachol stimulation (Fig. 9 E). The response to 1 mM carbachol was partially blocked by up to 1 nM RGS4 (Fig. 9 F), which converted the sustained response to [Ca2+]i oscillations. Complete inhibition of the response to 1 mM carbachol in Homer2−/− cells was observed at RGS4 concentrations above 2.5 nM (n = 4).
To corroborate this unexpected consequence of Homer 2 deletion, we measured the effect of recombinant Homer 2 on the GAP activity of RGS4 in an in vitro reconstitution system. The reconstitution system included the M1 muscarinic receptor and the heterotrimer Gαqβγ reconstituted into liposomes (Biddlecome et al., 1996). The Gαq GTPase activity in this system is dependent on receptor stimulation by carbachol and is inhibited by the muscarinic receptor antagonist atropine (Zeng et al., 1998). Fig. 10
C shows that Homer 2 increased RGS4 GAP activity by ∼90%. Importantly, stimulation of RGS4 GAP activity was specific to Homer 2, as Homer 1 had no effect.
PLCβ also functions as a Gα GAP (Cook et al., 2000). Because of the PIP2 hydrolyzing activity of PLCβ, it is not presently feasible to evaluate the significance of PLCβ GAP in vivo. However, to test the possibility that PLCβ GAP contributes to the enhanced signaling in Homer2−/− cells, we examined if Homers associate with PLCβ in vivo. For this we used GST-Homer 2, GST-Homer 1a and 1c, and two GST-Homer 1a mutants whose EVH domains were unable to bind PLCβ. Fig. 10 A shows that Homer 2 bound PLCβ in pancreatic acinar cell extract better than Homer 1c, and that Homer-PLCβ binding was inhibited by point mutations in the EVH domain. This point mutation destroys binding of all Homer isoforms. The general applicability of these findings is demonstrated by showing a similar profile of Homer-PLCβ binding in brain extract (Fig. 10 B). Homers 1 and 2 bound a comparable amount of mGluR5 monomers and dimers from the same brain extract, and this binding was abolished by the mutations in the EVH domain. The same experiment could not be done with RGS proteins due to a lack of antibodies with sufficient affinity to detect the low level of native RGS proteins.
The preferential binding of PLCβ to signaling complexes containing Homer 2 prompted us to test the effect of Homer 2 on PLCβ-GAP activity in the in vitro reconstitution system. Fig. 10 (C and D) shows the effect of Homer 2 on both PLCβ GAP activity and PIP2 hydrolysis. Homer 2 stimulated PLCβ GAP activity by ∼140%. This was better than Homer 2 stimulation of RGS4 GAP activity. Like RGS4 stimulation, PLCβ stimulation was specific for Homer 2; Homer 1 had no effect on PLCβ GAP activity. Homer 2 stimulated PLCβ PIP2 hydrolytic activity by only 35%. Homer 2 regulation of RGS4 and PLCβ GAP explains at least in part the effect of Homer 2 deletion on Ca2+ signaling.
The findings in the present work reveal several unexpected novel functions of Homer proteins and clarify the role of Homer 2 in Ca2+ signaling. The first finding of note is that expression of Homer proteins is isoform specific. Homer 3 is expressed in the basal pole, whereas Homers 1 and 2 are restricted to the apical pole. This already indicates that Homers must have nonoverlapping cellular functions. Indeed, Homer 3 has no discernible function in Ca2+ signaling in pancreatic acinar cells. Second, based on the modular structure of Homer proteins (EVH and C-C domains and a leucine zipper; Fagni et al., 2002) and their ability to bind the GPCR mGlu5 and IP3Rs (Tu et al., 1998), it is generally assumed that Homer proteins act as scaffolding proteins to assemble Ca2+-signaling complexes in cellular microdomains (Hering and Sheng, 2001; Fagni et al., 2002). Our results show that this is not the case for either Homer 2 or 3. Thus, deletion of these Homers (and of all Homers together) does not affect localization or expression of any IP3R isoform. The only effect observed is increased SERCA2b protein and activity in Homer2−/− cells. This is probably due to an adaptive response of the Ca2+-signaling system to the increased responsiveness of Homer2−/− cells to agonist stimulation. Several previous papers in heterologous systems (Liu et al., 1996; Brini et al., 2000) and in vivo (Zhao et al., 2001) have shown translational adaptation of the Ca2+-transporting machinery to disturbances in expression of Ca2+-transporting proteins.
The most notable finding in the present work is that Homer 2 functions to tune the intensity of agonist stimulation through regulation of RGS proteins and PLCβ GAP activities. Thus, deletion of Homer 2 left-shift the dose response for agonist-stimulated IP3 production and Ca2+ signaling without affecting the activation of PLCβ by activated Gα, while reducing the effectiveness of exogenous RGS proteins to inhibit Ca2+ signaling in vivo. These findings were corroborated by demonstrating direct activation of RGS proteins and PLCβ GAP activities in reconstitution system with recombinant proteins. The physiological significance of these findings is demonstrated by the nearly twofold increased frequency of [Ca2+]i oscillations in Homer2−/− cells.
In a previous work, we showed that RGS proteins provide a biochemical control of [Ca2+]i oscillations (Luo et al., 2001), thus, providing a molecular mechanism for oscillatory changes in [IP3] (Hirose et al., 1999; Nash et al., 2001) to drive [Ca2+]i oscillations. This mechanism explicitly implies that the activity of RGS proteins oscillates to induce oscillations in [IP3] and, consequently, [Ca2+]i oscillations (Luo et al., 2001). How the activity of RGS proteins is regulated during [Ca2+]i oscillations remains unclear. Here, we show that Homer 2 can regulate this activity in vivo and in vitro. Furthermore, we show that PLCβ GAP activity is not fixed, but is also regulated by Homer 2. Based on these findings, we propose that Homer 2 participates in regulating the GTPase reaction in the G protein turnover cycle to tune stimulus intensity.
At present, it is not clear how Homer 2 regulates RGS proteins and PLCβ GAP activities. However, the findings that Homer 2 preferentially binds PLCβ in pancreatic and brain extracts, that stimulation of GAP activity by Homer 2 can be reproduced in the minimal in vitro system, that Homer 2 activates GAP activity of two very different proteins whose only common feature is that they can bind to Gαq, and that Homer 2 has protein–protein binding domains all suggest that Homer 2 may control the proximity of the GAPs to Gαq and, thus, the efficiency of the GTPase reaction. This possibility and the molecular details of how Homer 2 controls GAP activity in vivo remain to be determined.
Regulation of RGS proteins and PLCβ GAP activities by Homer 2 raises the question of the significance of Homer binding to GPCRs and IP3Rs in Ca2+ signaling. Binding of Homer 2 or any of the Homers to one protein does not exclude binding and regulation of other proteins within the Ca2+-signaling complex. A more intriguing possibility is that different Homer isoforms mediate each of the interactions. In either case, it is clear that although Homer proteins do not appear to be the central scaffolding proteins that assemble the Ca2+-signaling complexes, they play an essential role in controlling Ca2+ signaling. One of their central roles is tuning Ca2+-signaling intensity. By tuning signal intensity, Homer 2 determines the frequency of [Ca2+]i oscillations and, in this way, controls the many cellular functions regulated by [Ca2+]i oscillations (Carafoli, 2002).
Materials And Methods
Cyclopiazonic acid, 1,4,5 and 2,4,5 IP3 were purchased from Qbiogene. The antibodies used in the present work were obtained from the following sources. Anti-Homer polyclonal antibodies (pAbs), and Homers 1, 2, and 3 were prepared as described previously (Tu et al., 1998). Anti-IP3R1, 2, and 3 pAbs were a gift from Dr. A. Tanimura (University of Hokkaido, Ishikari-Tobetsu, Japan). Anti-IP3R3 mAb was purchased from Transduction Laboratories. Anti-PMCA mAb was purchased from Affinity BioReagents, Inc. Anti-SERCA2b pAb was a gift from Dr. F. Wuytack (Catholic University of Leuven, Leuven, Belgium). Purified RGS4 and PLCβ were prepared as described previously (Zeng et al., 1998).
Generation of Homer 2 and 3 mutant mice
The homer 2 targeting construct was generated by inserting transprimer-1 into exon 3 of a 10.2-kb BAC fragment of the mouse homer 2 gene (Fig. 1). PGK-Neo was cloned into the unique PmeI site, and the resulting vector was subcloned into an MC1-TK vector. The resulting targeting construct was linearized and electroporated into R1 ES cells. Cells were selected with G-418 and gancyclovir. Clones were picked, screened by PCR, and confirmed by Southern blotting for homologous recombination. Clones were injected into blastocysts, and chimeras were mated to C57BL/6 mice to produce Homer 2 heterozygotes that were crossed to generate WT and Homer2−/− mice. Homer3−/− mice were generated using a similar strategy as that used to delete Homer2, except that the PGK-Neo cassette was directly inserted into exon 3 of Homer3 at the SmaI site.
Preparation of pancreatic acini
Acini were prepared from the pancreas of WT, Homer2−/−, and Homer3−/− mice by limited collagenase digestion as described previously (Shin et al., 2001). After isolation, the acini were resuspended in a standard solution A ([mM] 140 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 10 Hepes, pH 7.4 with NaOH, 10 glucose, and 0.1% BSA), and kept on ice until use. Doublet or triplet acinar cell clusters were obtained by incubating a minced pancreas in a 0.025% trypsin, 0.02% EDTA solution for 5 min at 37°C. After washing with solution A supplemented with 0.02% soybean trypsin inhibitor (PSA), doublets and triplets were liberated by a 7-min incubation at 37°C in the same solution that also contained 160 U/ml pure collagenase. The cells were washed with solution A and kept on ice until use.
Measurement of [Ca2+]
Cells in PSA were incubated with 5 μM Fura2/AM for 30 min at room temperature and washed once with PSA. Samples of cells were plated on glass coverslips that formed the bottom of a perfusion chamber. After 2–3 min of incubation, to allow cell attachment to the coverslip, the cells were continuously perfused with prewarmed (37°C) solution A at a rate of 5 ml/min (30 vol chamber/min). Agonists were delivered to the cells by inclusion in the perfusate. Fura2 fluorescence was measured at excitation wavelengths of 340 and 380 nm using a PTI image acquisition and analysis system as detailed previously (Shin et al., 2001).
The whole cell configuration of the patch clamp technique was used for measurement of Ca2+-activated Cl− current as a reporter of [Ca2+]i next to the PMCA. The experiments were performed with single acinar cells perfused with solution A. The standard pipette solution contained (mM): 140 KCl, 1 MgCl2, 0.1 EGTA, 5 ATP, 10 Hepes (pH 7.3 with KOH) with or without 2,4,5 IP3 or between 0.25 and 10 nM RGS4, as described previously (Zeng et al., 1996). The RGS4 was dialyzed against an ATP-free pipette solution and concentrated to ∼5 μM with a centricone system. Seals of 6–10 GΩ were produced on the cell membrane, and the whole cell configuration was obtained by gentle suction or voltage pulses of 0.5 V for 0.3–1 ms. The patch clamp output (Axopatch-1B; Axon Instruments, Inc.) was filtered at 20 Hz. Recording was performed with patch clamp 6.0 and a Digi-Data interface (model 1200; Axon Instruments, Inc.). The current was recorded at a holding potential of −40 mV. The oscillation frequency was determined from a stretch of at least 5 min starting at the first full Ca2+ spike. The number of spikes over this time period was counted to determine the number of spikes/minute and results from at least five cells from two mice were used to obtain the results listed in the text and shown in Fig. 7.
Measurement of Ca2+ uptake and release from internal stores
IP3-mediated Ca2+ release from internal stores was measured in SLO-permeabilized cells as described previously (Xu et al., 1996b). In brief, cells washed with a high K+, Chelex-treated medium were added to the same medium containing an ATP regeneration system (comprised of 3 mM ATP, 5 mM MgCl2, 10 mM creatine phosphate, and 5 U/ml creatine kinase), a cocktail of mitochondrial inhibitors, 2 μM Fluo3 and 3 mg/ml SLO (Difco). In this medium, the cells were almost instantaneously permeabilized so that Ca2+ uptake into the ER could be measured immediately. Uptake of Ca2+ into the ER was allowed to continue until medium [Ca2+] was stabilized. Then IP3 was added in increasing concentrations to measure the extent of Ca2+ release and the potency of IP3 in mobilizing Ca2+ from the ER.
Measurement of IP3 mass
IP3 levels were measured by a radioligand assay (Xu et al., 1996b). Intact acini were stimulated with the indicated concentration of agonist for 5 s at 37°C. Permeabilized acini were stimulated with 2.5–100 μM GTPγS for 15 s at 37°C. Preliminary experiments showed that 1,4,5 IP3 reached maximal level at these incubation times. The reactions were stopped by the addition of 100 μl ice-cold 15% PCA to 100 μl of samples, vigorous mixing, and incubation on ice for at least 10 min to allow precipitation of proteins. Standards of IP3 were prepared in the same manner. The perchloric acid was removed, and IP3 was extracted with 0.2 ml Freon and 0.2 ml tri-n-octylamine. IP3 content in the aqueous phase was measured by displacement of [3H]IP3 using microsomes prepared from bovine brain cerebella.
Microsomes were prepared by homogenizing pancreatic acini or brains from WT, Homer2−/−, and Homer3−/− mice in a buffer containing (mM): 100 KCl , 20 Tris-base, pH 7.6 with KOH, 1 EDTA, 1 benzamidine, and 1 PMSF. The homogenates were centrifuged at 1,000 g for 10 min at 4°C. The supernatants were collected and centrifuged at 40,000 g for 30 min. The pellets were resuspended in homogenization buffer and the microsomes were stored at −80°C until use. Microsomes were extracted by a 1-h incubation on ice in a buffer containing (mM): 50 Tris, pH 6.8 with HCl, 150 NaCl, 2 EDTA, 2 EGTA, and 1% Triton X-100 supplemented with protease inhibitors (0.2 mM PMSF, 10 μg/ml leupeptin, 15 μg/ml aprotinin, and 1 mM benzamidine). The extracts were used to separate the proteins by SDS-PAGE and the proteins were probed with a 1:500 dilution of SERCA2b pAb; 1:500 dilution of IP3R1 pAb; and 1:1,000 dilution of IP3R3 and PMCA mAbs.
Rat pancreatic acini were used to prepare extracts as described above. Adult rat forebrain was sonicated in four volumes of ice cold PBS with 1% Triton X-100. Soluble extracts were mixed with GST-Homer proteins lined to glutathione beads (Sigma-Aldrich) for 2 h (pancreas) or overnight (brain) at 4°C. Beads were pelleted and washed four times in lysis buffer and eluted with loading buffer followed by SDS-PAGE. The blots were probed for PLCβ, mGlu5α, or stained with Coomassie blue as before (Tu et al., 1998; Xiao et al., 2000).
Cells from WT, Homer2−/−, and Homer3−/− mice attached to glass coverslips were fixed and permeabilized with 0.5 ml of cold methanol for 10 min at −20°C, except for the experiments in Figs. 2 and 4 for localization of Homer 3, in which the cells were fixed with 4% formaldehyde for 20 min at room temperature, followed by permeabilization with 0.05% Triton X-100. After removal of methanol or Triton X-100, the cells were washed with PBS and incubated in 0.5 ml PBS containing 50 mM glycine for 10 min at room temperature. This buffer was aspirated and the nonspecific sites were blocked by a 1-h incubation at room temperature with 0.25 ml PBS containing 5% goat serum, 1% BSA, and 0.1% gelatin (blocking medium). The medium was aspirated and replaced with 50 μl of blocking medium containing control serum or a 1:50 dilution of pAbs against Homer 1, 2, or 3, or a 1:100 dilution of pAb against IP3R1, 2, or 3. After incubation with the primary antibodies overnight at 4°C and three washes with the incubation buffer (same as blocking buffer, but without serum), the antibodies were detected with goat anti–rabbit or anti–mouse IgG tagged with fluorescein or rhodamine. Images were collected with a confocal microscope (model MRC 1024; Bio-Rad Laboratories).
Measurement of GTPase and PLC activities in vitro
Agonist-stimulated steady-state GTP hydrolysis was measured in reconstituted vesicles that contained recombinant, purified heterotrimeric Gq and M1 acetylcholine receptors at 30°C in the presence and absence of 2–5 nM RGS4 or PLCβ1 (Biddlecome et al., 1996). Homers 1 and 2 (250 nM) were preincubated with the vesicles in the presence of 1 mM carbachol and one of the GAPs for 45 min at 0°C followed by further incubation for 2 min at 30°C (Mukhopadhyay and Ross, 1999). The GTPase reaction was initiated by adding 3 μM γ-[32P]GTP and was continued for 10 min at 30°C. Reactions were quenched with a cold slurry of Norit in H3PO4 and [32P]Pi was measured in the supernatant.
The effect of Homer 2 on steady-state phospholipase activity of PLCβ1 was measured with the same reconstituted phospholipid vesicles, which also contained [3H]PIP2. Homer 2 was preincubated with PLCβ1 for 45 min at 0°C. PIP2 hydrolysis was initiated by adding a mixture of the preincubated Homer 2 (250 nM final) and PLCβ1 (2 nM final) to the reconstituted vesicles. [3H]IP3 release was measured at 30°C in the presence of 1 mM carbachol, 10 μM GTP, and 10 nM-free Ca2+ as described previously (Biddlecome et al., 1996).
When appropriate, results are given as the mean ± SEM of the indicated number of experiments. Statistical significance was evaluated by a two-way ANOVA. All immunostaining experiments were repeated at least five times with similar results.
We thank Jin Chen for superb technical assistance and Daniella Muallem for help in editing the manuscript.
This work was supported by grants from the National Institute of Diabetes and Digestive and Kidney Diseases, the National Institute on Deafness and Other Communication Disorders, the National Institute of General Medical Sciences, the National Institute of Drug Abuse, and the National Institute of Mental Health. Support was also provided by a Korea Research Foundation Grant (KRF-2002-015-EP0115) to D.M. Shin.
D.M. Shin, M. Dehoff, and X. Luo contributed equally to this work.